Spatiotemporal regulation of nervous system development in the annelid Capitella teleta
© The Author(s) 2017
Received: 8 June 2017
Accepted: 20 July 2017
Published: 1 August 2017
How nervous systems evolved remains an unresolved question. Previous studies in vertebrates and arthropods revealed that homologous genes regulate important neurogenic processes such as cell proliferation and differentiation. However, the mechanisms through which such homologs regulate neurogenesis across different bilaterian clades are variable, making inferences about nervous system evolution difficult. A better understanding of neurogenesis in the third major bilaterian clade, Spiralia, would greatly contribute to our ability to deduce the ancestral mechanism of neurogenesis.
Using whole-mount in situ hybridization, we examined spatiotemporal gene expression for homologs of soxB, musashi, prospero, achaete–scute, neurogenin, and neuroD in embryos and larvae of the spiralian annelid Capitella teleta, which has a central nervous system (CNS) comprising a brain and ventral nerve cord. For all homologs examined, we found expression in the neuroectoderm and/or CNS during neurogenesis. Furthermore, the onset of expression and localization within the developing neural tissue for each of these genes indicates putative roles in separate phases of neurogenesis, e.g., in neural precursor cells (NPCs) versus in cells that have exited the cell cycle. Ct-soxB1, Ct-soxB, and Ct-ngn are the earliest genes expressed in surface cells in the anterior and ventral neuroectoderm, while Ct-ash1 expression initiates slightly later in surface neuroectoderm. Ct-pros is expressed in single cells in neural and non-neural ectoderm, while Ct-msi and Ct-neuroD are localized to differentiating neural cells in the brain and ventral nerve cord.
These results suggest that the genes investigated in this article are involved in a neurogenic gene regulatory network in C. teleta. We propose that Ct-SoxB1, Ct-SoxB, and Ct-Ngn are involved in maintaining NPCs in a proliferative state. Ct-Pros may function in division of NPCs, Ct-Ash1 may promote cell cycle exit and ingression of NPC daughter cells, and Ct-NeuroD and Ct-Msi may control neuronal differentiation. Our results support the idea of a common genetic toolkit driving neural development whose molecular architecture has been rearranged within and across clades during evolution. Future functional studies should help elucidate the role of these homologs during C. teleta neurogenesis and identify which aspects of bilaterian neurogenesis may have been ancestral or were derived within Spiralia.
KeywordsNeurogenesis Annelid Capitella teleta Spiralian SoxB1 Neurogenin Ash NeuroD Musashi Prospero
Neurogenesis refers to the process by which differentiated neurons are generated from neural precursor cells (NPCs), and a critical component of this process is to maintain a balance between cell proliferation and differentiation. Coordination between these two distinct processes is necessary for differential growth and generation of a variety of functional neural cell types at the correct time and place, and an imbalance can result in harmful developmental defects [1, 2]. Various signaling pathways, transcription factors, and RNA-binding proteins regulate neurogenesis, and alterations in neurogenic regulatory networks and patterns of gene expression underlie the evolution of many complex phenotypes, including nervous systems [3–6]. Therefore, studying gene expression of neurogenic factors across a broad range of taxa can help us understand evolution of the molecular mechanisms underlying nervous system development.
Molecular and cellular studies have revealed that vertebrates and insects share some neurogenic mechanisms but there are also differences [1, 7–10]. One commonality between arthropods and vertebrates is that dedicated NPCs generate the various neural cell subtypes by asymmetric cell division [7, 11]. Some of the key molecular regulators of neurogenesis include SoxB, basic helix-loop-helix (bHLH) group A transcription factors (including Achaete–Scute complex a or ASCa family members), and Notch signaling. In vertebrates, SoxB1 homologs (e.g., mouse Sox1, 2, and 3) are initially expressed throughout the neuroectoderm and then in mitotically active NPCs. SoxB1 expression is largely downregulated as NPCs begin to differentiate [12–16]. Vertebrate NPCs rely on SoxB1 transcription factors and Notch signaling to remain in a proliferative, undifferentiated state. In part, SoxB1 and Notch signaling maintain NPCs by reducing expression levels and activity of proneural bHLH transcription factors such as Neurogenin1 and 2 and Mash1 (an ASCa homolog). Proneural proteins in turn promote cell cycle exit and neuronal differentiation, repress SoxB1 activity, and upregulate expression of the Notch ligand delta [9, 12, 13, 17–21]. One mechanism by which proneural proteins suppress SoxB1 activity is by upregulating expression of the SoxB2 gene Sox21, which promotes neural differentiation .
The functions of the D. melanogaster SoxB homologs SoxNeuro and Dichaete (also known as Fish-hook) are similar to vertebrates in that they help maintain neuroblasts [20, 23]. SoxNeuro is expressed throughout the neuroectoderm but gets downregulated in delaminated neuroblasts [24–26], while Dichaete has a somewhat more dynamic expression pattern in the neuroectoderm and neuroblasts [27, 28]. Loss of function of SoxNeuro and Dichaete results in a loss of neuroblasts throughout the neuroectoderm and severe hypoplasia in the CNS [24, 26, 29]. Proneural bHLH factors in the ASCa family, particularly Achaete, Scute, and Lethal of Scute, are also involved in neurogenesis in insects. However, they have a slightly different function than in vertebrates—they promote fate specification of neuroblasts at the expense of epidermal stem cells. ASCa proteins upregulate delta expression in presumptive neuroblasts, and then Delta activates Notch on neighboring cells. Targets of activated Notch downregulate expression of ASCa genes, thus preventing cells from becoming neuroblasts [9, 30–32]. There is evidence that the SoxB proteins in D. melanogaster can directly regulate gene expression of achaete and asense [33–36]. However, it is not clear whether proneural bHLH proteins affect expression of soxB homologs as they do in vertebrates, and definitive SoxB2 homologs have not yet been identified in D. melanogaster [37, 38].
Differences in neurogenic mechanisms can also be seen within clades. For example, in earlier branching arthropods such as the spider Cupiennius salei and the myriapods Glomeris marginata and Lithobius forficatus, ASCa homologs (ash) are expressed in patches of mitotically quiescent neuroectodermal cells, which separate from the apical surface as groups and eventually differentiate into neural cells [39–43]. This contrasts with insects, where proneural gene expression initiates within each proneural cluster but then becomes limited to the neuroblast, which will go on to divide [44, 45]. Within select clades of Spiralia (e.g., Platyhelminthes, annelids, mollusks, and nemerteans), recent studies highlight a potential involvement of SoxB and proneural proteins during neurogenesis [46–49]. In the annelid Platynereis dumerilii, Pdu-ASH and Pdu-Ngn are expressed along the apical proliferating zone of the neuroectoderm, while Pdu-SoxB is expressed throughout the neuroectoderm at earlier stages [48, 49]. Such variation highlights the importance of studying neurogenesis in multiple species within clades in order to understand what aspects of bilaterian neurogenesis are ancestral and what aspects have been derived within particular taxa. Our understanding of neurogenesis in spiralians, including the molecular components, the exact role of each component, and the extent of variability in the molecular and cellular details of neurogenesis in this clade is still relatively incomplete. Furthermore, a proper understanding of neurogenesis in spiralians is required to reconstruct the evolution of nervous systems within Bilateria.
In this article, we extend previous studies to describe the spatiotemporal expression of candidate neurogenic genes in the annelid Capitella teleta. Some aspects of neurogenesis have previously been described in C. teleta . However, neurogenic mechanisms underlying ventral nerve cord (VNC) development, including gene expression, have not yet been well characterized. We found that gene homologs of SoxB, Musashi (Msi), Prospero (Pros), Achaete–Scute (Ash), Neurogenin (Ngn), and NeuroD are expressed in developing neural tissue in C. teleta. The onset and duration of expression and spatial localization indicate roles during different phases of neurogenesis within the brain and VNC.
Capitella teleta  adults were maintained in the laboratory as previously described [52, 53]. Animals were kept in bowls of artificial seawater (ASW) and mud at 19 °C. Every 2 weeks, the adult worms were transferred to new bowls in order to maintain the density of worms within each bowl. Broods were dissected using a clean pair of Dumont #5 forceps to release the different embryonic and larval stages reared by the females. Embryonic and larval stages were collected from different bowls and used for whole-mount in situ hybridization (WMISH) experiments.
Isolation of C. teleta neurogenic gene homologs
Total RNA was extracted from mixed stage 1–9 embryos and larvae using the RNA Trizol extraction protocol (Molecular Research Center, Inc.) or the RNeasy Mini Kit (Qiagen). Reverse transcription reactions were conducted using the SMARTer RACE kit (Clontech). Capitella teleta homologs were identified by tBLASTn searches against the C. teleta genome and EST libraries (JGI, DOE). We identified two soxB orthologs and single orthologs of musashi, prospero, neurogenin, and neuroD. We named these genes Ct-soxB1, Ct-soxB, Ct-msi, Ct-pros, Ct-ngn, and Ct-neuroD, respectively. Fragments of the coding sequences of these genes were amplified by PCR using gene-specific primers as follows.
Ct-soxB1 (GenBank accession # MF508645): 917 bp containing some 5′ UTR, an HMG box, and a partial Soxp domain; Fwd primer 5′-CAAAGTCCTCGCTCAAAGCAG and Rev primer 5′-GCATGTATCCGTTCATGTTCATAGAG.
Ct-soxB (MF508646): 740 bp containing some 5′ UTR, an HMG box, and a partial Soxp domain; Fwd primer 5′-TTACCCTTCAACAAATCTAACTGC and Rev primer 5′-CGTATGGCGAGTAGAAAGCTC.
Ct-msi (MF508642): 729 bp containing a majority of the coding domain, including both RNA recognition motifs (RRMs), and some 3′-UTR; Fwd primer 5′-AGCCAGCAATCTACGTCAGG; Rev primer 5′-CACCACAGCAACGTGTTACC.
Ct-prospero (MF508647): 711 bp containing some 5′ UTR, the open reading frame with the entire homeo-prospero domain (HPD), and some 3′-UTR; Fwd primer 5′-CCAAGAACAGAAAAAGCAC; Rev primer 5′-TGTTTTGACTGCTTGATA.
Ct-neurogenin (MF508643): 778 bp of open reading frame with the entire bHLH domain; Fwd primer 5′-GGTCAATCTGACAGCAAGCA; Rev primer 5′-GGTAATGTCCTTGGTAACCTGGC.
Ct-neuroD (MF508644): 669 bp of open reading frame with the entire neuronal bHLH domain; Fwd primer 5′-ATGGCTAAAGCAGGAGATG; Rev primer 5′-TTCGGGTGATAGCGAGTAGG.
PCR products were TA cloned into the pGEM-T Easy vector (Promega) and sequenced. These gene fragments were used as templates to generate DIG-labeled anti-sense RNA probes for WMISH. A fragment of Ct-ash1 was previously clones and is described in .
Gene orthology analyses
Amino acid sequences for Sox family proteins (Additional file 1: Figure S1) and Musashi family proteins (Additional file 2: Figure S2) were aligned in MacVector 15.1 (MacVector, Inc.) via the T-Coffee alignment tool. The alignments were trimmed using trimAl v1.2b  via the gappyout algorithm. The best-fit model of protein evolution for each alignment was determined via ProtTest v3.2 [55, 56]. Bayesian phylogenetic analyses were performed on these alignments with MrBayes v3.2  using the LG+G (Sox) or LG+G+I (Musashi) models of protein evolution  with 1,000,000 generations sampled every 100 generations and four chains. Fifty percent majority-rule consensus trees were produced from the last 7500 trees of each analysis representing 750,000 stationary generations. Consensus trees were visualized with FigTree v1.4.3 . Posterior probabilities and branch lengths were calculated from these consensus trees. Maximum likelihood phylogenetic analyses were performed on these alignments with RAxML v8.2.10  using the LG+G model of protein evolution and rapid bootstrapping with 500 replicates. Consensus trees were produced using the web-based interactive tree of life (iTOL) v3.5 . Branches with a bootstrap value below 45 were collapsed.
In situ hybridization
Multiple animals of each stage were obtained from different broods and treated for fixation as described elsewhere [50, 52]. All fixations for WMISH were done in 4% paraformaldehyde in ASW for more than 6 h. After fixation, animals were serially dehydrated in methanol and stored at −20 °C. WMISH was conducted as described previously . Briefly, animals were hybridized for a minimum of 72 h at 65 °C with 1 ng/µl of each probe. DIG-labeled RNA probes were generated using the MegaScript SP6 or T7 transcription kit (ThermoFisher Scientific). Spatiotemporal RNA localization was observed using an NBT/BCIP color reaction. After WMISH, animals were stained with 0.1 µg/ml Hoechst 33342, cleared in 80% glycerol in PBS, and mounted on microscope slides for DIC imaging.
Images were taken using DIC optics on either a (1) Zeiss M2 microscope with an 18.0 megapixel EOS Rebel T2i digital camera (Canon) or (2) a Zeiss Axioskop 2 microscope and a SPOT Flex digital camera (Diagnostic Instruments, Inc, MI, USA) in conjunction with the Spot Advanced version 4.6 software (Diagnostic Instruments, Inc). Animals from multiple broods were imaged for each developmental stage and gene. WMISH images were cropped and brightness, contrast, and color balance edited to maintain consistency within plates using Adobe Photoshop CC. Figure panels were constructed using Adobe Illustrator CC.
Overview of larval neurogenesis in Capitella teleta
The VNC in C. teleta develops from anterior to posterior, and cell division occurs throughout the ventral neuroectoderm from stages 4–6 . Toward the end of stage 6, cell division begins to be restricted to the posterior end of the larva and by stage 7 largely takes place in the posterior growth zone (PGZ; Fig. 1b, pink) . Furthermore, by stage 7, mature neurons are present in all 13 ganglia formed in larvae as demonstrated by expression of the pan-neuronal marker Ct-syt1 and the presence of neurons with FMRF-like immunoreactivity [50, 52]. By stage 8, cell division in the trunk has subsided except in the PGZ , suggesting that neurogenesis in the first 13 ganglia is complete. Ganglia continue to be added to the VNC from the PGZ as new segments form in juveniles and adults .
Ct-soxB1 and Ct-soxB expression
Description of Ct-soxB1: Ct-soxB1 transcript was detected in zygotes and early cleavage stages (two cells through late cleavage; data not shown). Once the embryos begin to gastrulate (stage 3), Ct-soxB1 is expressed in the anterior ectodermal thickening (Fig. 2a–d, arrow), which comprises brain NPCs . Ct-soxB1 is also present at lower levels in a majority of micromeres during gastrulation (data not shown), but becomes downregulated in most cells outside of the anterior ectoderm by the end of stage 3 (Fig. 2b–d). After gastrulation is complete (stage 4), Ct-soxB1 continues to be expressed throughout the anterior ectoderm, but is more highly expressed in the neuroectoderm (Fig. 2e, f, h). At stage 4, Ct-soxB1 transcript also is detected in the trunk in the presumptive neuroectoderm of the VNC (ventral neuroectoderm; Fig. 2g) and in patches of cells dispersed throughout the pygidial ectoderm (Fig. 2h, black arrowhead points to one cell). Furthermore, the onset of Ct-soxB1 expression in the ventral neuroectoderm precedes formation of ganglia and expression of neural differentiation markers such as Ct-syt1 by at least one full stage, or 1 day at 19 °C .
During stage 5, Ct-soxB1 is expressed in surface neuroectoderm, where cells are dividing, as well as in the majority of cells that have already internalized to form the brain (Fig. 2i, j, l). In the trunk, the expression of Ct-soxB1 expression expands considerably, extending from the mouth to the telotroch (Fig. 2k) and covering the entire ventrolateral domain (data not shown). Ct-soxB1 is still expressed in the pygidial ectoderm, as well as in two new domains, in the foregut (Fig. 2k, l) and dorsolateral ectoderm (data not shown). At stage 6, Ct-soxB1 continues to be expressed in the surface neuroectoderm in the episphere as well as in cells in the developing brain (Fig. 2m, n, p), although expression appears to be excluded from the most basal portion of the brain (Fig. 2n, dashed line demarcates the basal edge of the brain). Ct-soxB1 is also present in individual ectodermal cells that form a circumferential ring immediately posterior to the prototroch (Fig. 2o, black arrowhead). Within the trunk at stage 6, a subset of ventral neuroectodermal cells express Ct-soxB1 as well as individual cells within the developing VNC (Fig. 2o, arrowheads; Fig. 2p). At this stage, lower levels of Ct-soxB1 expression are still detected in the developing foregut and the lateral and dorsolateral ectoderm (Fig. 2o, p and data not shown).
By stage 7, as larval neurogenesis in the episphere is nearing completion, Ct-soxB1 is expressed in the brain and the overlying epidermis (Fig. 2q, s). There also is a new region of single cells in the ventral epidermis in the episphere (Fig. 2r, arrows) and adjacent to the opening of the mouth (Fig. 2r). Within the trunk, Ct-soxB1 continues to be expressed in a circumferential row of epidermal cells posterior to the prototroch (Fig. 2r, black arrowhead). Ct-soxB1 is expressed in a small subset of cells in the VNC as well as in six longitudinal rows of ectodermal cells (three rows on either side of the body), which are likely part of the peripheral nervous system. The ventral-most pair of longitudinal rows is positioned on either side of the VNC (Fig. 2r, white arrowheads). The second pair of longitudinal rows is also ventrolateral, while the third pair of rows is dorsolateral (data not shown). Trunk expression also persists in the foregut (Fig. 2s) and can be seen in the PGZ, where new segments and ganglia of the VNC are being added (Fig. 2s, arrow). The pattern of expression at stage 8 is very similar to that observed at stage 7 (Additional file 4: Figure S4a–c; arrowhead in a points to expression the VNC; arrow in c points to expression in the epidermis in the episphere). Moreover, the Ct-soxB1 + cells in the ventral epidermis are quite distinct at stage 8, forming a ‘grid-like’ pattern (Additional file 4: Figure S4a). By stage 9, Ct-soxB1 is downregulated in most of the larva except for a few cells in the epidermis of the episphere and in the PGZ (Additional file 4: Figure S4d).
Description of Ct-soxB: Ct-soxB was detected in zygotes and early cleavage stage embryos (two cells through birth of the third quartet of micromeres; data not shown). Once gastrulation begins, Ct-soxB is expressed in the anterior ectodermal thickening (Fig. 3a, arrow). In later stage 3 embryos, Ct-soxB is expressed in patches of cells throughout the anterior ectoderm, with higher levels in the anterior ectodermal thickening (Fig. 3b, c, arrow), which is similar to Ct-soxB1 at this stage. Ct-soxB is also expressed in a circumferential band of cells in the anterior half of the trunk (data not shown). During stage 4, Ct-soxB continues to be expressed in the anterior ectoderm, and expression in the neuroectoderm appears to be confined to surface cells (Fig. 3d–f). By the end of stage 4, Ct-soxB expression also becomes visible in cells around the stomodeum (data not shown) and in the pygidial ectoderm, posterior to the telotroch, (Fig. 3e, f, arrows). Unlike Ct-soxB1 at stage 4, Ct-soxB does not appear to be expressed in the ventral neuroectoderm at this stage.
During stage 5, as the brain lobes become more visible, Ct-soxB continues to be expressed in surface anterior neuroectoderm and also begins to be expressed and at varying intensities throughout the developing brain (Fig. 3g, h, j). In the trunk, Ct-soxB is expressed around the stomodeum and in patches within the ventral neuroectoderm (Fig. 3i). Pygidial expression continued to be detected at stage 5 (Fig. 3j), but not thereafter. At stage 6, Ct-soxB continues to be expressed in patches of cells in surface anterior neuroectoderm and in the developing brain (Fig. 3k, l, p). Expression in the trunk at stage 6 encompasses cells around the stomodeum and a small subset of cells in the VNC (Fig. 3m, p, arrowheads). In addition, Ct-soxB is expressed in two ventrolateral rows of cells on either side of the VNC (Fig. 3m–o white arrowheads) and in patches of cells laterally (Fig. 3n, o, arrows) and dorsolaterally (data not shown). There is also a circumferential row of Ct-soxB + cells immediately posterior to the prototroch (Fig. 3o, black arrowhead).
At stage 7, Ct-soxB is expressed in a ‘grid-like’ pattern in the ventral epidermis of the episphere and around the mouth opening (Additional file 4: Figure S4e), similar to Ct-soxB1. Additional expression domains include the brain lobes, the epidermis overlying the brain, and a subset of cells in the pharynx (Additional file 4: Figure S4f). Ct-soxB was also detected in a few cells in the developing VNC (Additional file 4: Figure S4e, arrowheads) as well as in the ventrolateral ectoderm (data not shown). Ct-soxB expression at stage 8 is similar to that at stage 7, except that expression in the VNC and ventrolateral ectoderm was not detected (Additional file 4: Figure S4g). By stage 9, expression was only detected in the brain (Additional file 4: Figure S4h).
Ct-msi homolog expression
Ct-msi transcripts were detected in zygotes and two-cell embryos through birth of the third quartet of micromeres, although expression was only detected in a few micromeres after birth of the third quartet (data not shown). During gastrulation, Ct-msi was detected in the anterior ectodermal thickening (Fig. 4a, b, arrow) as well as around the blastopore opening and in internalized endodermal cells (Fig. 4a, b, arrowhead). From late stage 3 through 4, Ct-msi is expressed in the anterior neuroectoderm (Fig. 4c–f, h), in cells abutting the stomodeum, and in the mesodermal bands (Fig. 4g, h; white arrowheads point to mesodermal staining). Two longitudinal rows of expression that may coincide with the developing VNC at this stage were also detected (Fig. 4g, h, black arrowheads).
Ct-msi continues to be expressed in the developing brain at stage 5 (Fig. 4i, j, l), and expression appears to be excluded from surface neuroectoderm (Fig. 4l, dashed line denotes the apical surface of the ectoderm). Similar to brain expression at this stage, Ct-msi is expressed in the developing VNC (Fig. 4k, m; black arrowheads in m point to expression in the VNC) and appears to be subsurface, although it is more difficult to discern surface versus subsurface for the VNC at this stage. Expression in the mesodermal bands (Fig. 4k, l, white arrowheads) and foregut (Fig. 4k–m) continues to be present at stage 5. Ct-msi also begins to be expressed in cells associated with the circumesophageal connectives (Fig. 4i, m arrows). At stage 6, Ct-msi expression persists throughout the developing brain (Fig. 4n, o, q) but is not present in the overlying ectoderm (Fig. 4q). Within the trunk, Ct-msi is expressed in the foregut and throughout the VNC (Fig. 4p, q). Interestingly, Ct-msi does not appear to be expressed in the surface ventral neuroectoderm (Fig. 4q, dashed line denotes the apical surface of the ectoderm). Expression in the mesoderm is greatly decreased at stage 6, although we did detect posteriorly localized, longitudinal rows of Ct-msi + cells (at least three rows on each side of the larva) positioned between the mesoderm and endoderm. These cells are positioned laterally and dorsally, and one dorsally localized row is indicated with a white arrowhead in the inset in Fig. 4q. At stages 7–9, Ct-msi continues to be expressed in the brain and VNC, but is not expressed in the mesoderm or surface ectoderm (Additional file 6: Figure S6a–e). We also did not detect Ct-msi transcript in the PGZ (Additional file 6: Figure S6a, b, d, e), where the majority of cell division occurs after stage 7 . Once the foregut has differentiated into a pharynx and esophagus (stage 8), it is clear that Ct-msi expression is restricted to the pharynx (Additional file 6: Figure S6d).
Ct-pros homolog expression
Ct-pros transcripts were detected in zygotes and in early cleavage stage embryos (two cells though birth of the third quartet of micromeres; data not shown). During early epiboly, Ct-pros is expressed in 3–4 cells at the animal pole (Fig. 5a, arrowheads). As the blastopore closes, a few Ct-pros + cells can be detected within the anterior neuroectoderm (Fig. 5b, arrowheads), in the region where the mouth will form, and in lateral and dorsal ectoderm (data not shown). In late stage 3 embryos, Ct-pros is expressed in single cells in the anterior ectoderm, including in the thickened region that will generate the brain (Fig. 5c, arrowheads). In the trunk, a circumferential ring of Ct-pros + cells can be detected in the ectoderm of the peristomium, at the anterior–posterior position of the mouth (Fig. 5d, arrow). Within the peristomium, there appear to be fewer Ct-pros + cells at the dorsal midline (data not shown). Finally, Ct-pros is expressed in single cells throughout the pygidial ectoderm (data not shown). At stage 4, Ct-pros is expressed in more cells in the anterior ectoderm and in subsurface cells in the developing brain (Fig. 5e, g). Expression of Ct-pros in the peristomial ectoderm (Fig. 5f, h) and the pygidial ectoderm (Fig. 5g) expands to encompass more cells, although there are still fewer Ct-pros + cells at the dorsal midline within the peristomium. Some animals also begin to express Ct-pros in a few lateral ectodermal cells just posterior to the peristomium (Fig. 5h, arrow).
At stage 5, Ct-pros continues to be expressed in the anterior ectoderm, brain, trunk ectoderm, and pygidial ectoderm (Fig. 5i–l). Additionally, at stage 5, Ct-pros is expressed in the developing VNC (Fig. 5k, l, black arrowheads). At stage 6, Ct-pros expression decreases to a subset of cells in anterior neuroectoderm and in the lateral and medial brain (Fig. 5m, n, q, r). Within the trunk at stage 6, Ct-pros is expressed in a subset of cells in the VNC (Fig. 5o, p, r, black arrowheads) and foregut (Fig. 5r) and in single cells throughout the ectoderm (Fig. 5o–q). Within the pygidium, Ct-pros is only expressed in the rectum (Fig. 5r, white arrowhead).
At stage 7, Ct-pros becomes further reduced in the episphere and is largely localized to the lateral sides of the brain (Additional file 6: Figure S6f, arrows). Within the trunk, Ct-pros is expressed in a subset of cells in the VNC (Additional file 6: Figure S6g, h, black arrowheads), in the foregut (data not shown), and in single cells in a circumferential band in the posterior ectoderm that encompasses the PGZ (Additional file 6: Figure S6g, bracket). Color product was also detected at the base of each chaeta at stage 7; however, this may be due to probe trapping rather than actual gene expression. Stage 8 and 9 animals also express Ct-pros along the lateral edges of the brain, in the pharynx and esophagus, in the VNC, and in the PGZ (Additional file 6: Figure S6i, j; black arrowheads denote expression in the VNC).
Ct-ngn homolog expression
Ct-ngn transcript was detected in zygotes through early cleavage stages (two cells through birth of the third quartet; data not shown). In stage 3 embryos undergoing epiboly, Ct-ngn is expressed on the animal side of the embryo in three patches, each patch comprising one to two cells (Fig. 6a, b, arrows and white arrow). Expression was also detected in a few smaller posteriolateral cells (data not shown). Two of the three patches of Ct-ngn + cells appear to be at the lateral edges of the anterior ectodermal thickening (Fig. 6a, b, arrows; anterior ectodermal thickening is bracketed in b), while the other Ct-ngn + patch of expression does not appear to be within the anterior ectodermal thickening (Fig. 6a, b, white arrow). As the blastopore begins to close, three to five single, large cells (usually four) in the anterior ectodermal thickening and a few smaller cells in the lateral and pygidial ectoderm express Ct-ngn (Fig. 6c; black arrowhead marks pygidial expression). By the end of stage 3, once the mouth opening is apparent, Ct-ngn expression is localized to a several single cells in the anterior neuroectoderm and the pygidium (Fig. 6d–f; black arrowhead indicates pygidial cells). The expression of Ct-ngn is similar from late stage 3 to early stage 4, with the addition of a few Ct-ngn + cells in the ventral neuroectoderm (Fig. 6k, arrowheads), in the ectoderm just posterior to the prototroch, and around the stomodeum, likely in the presumptive foregut (data not shown). At the end of stage 4, Ct-ngn expression in the anterior and ventral neuroectoderm expands (Fig. 6g–j), and new expression in a few dorsal and ventral cells in the ectoderm of the episphere can be detected (Fig. 6g). Pygidial expression of Ct-ngn was not detected in late stage 4 larvae (data not shown).
By stage 5, increasing numbers of superficial cells in the anterior neuroectoderm express Ct-ngn (Fig. 6l, m, o). The trunk expression expands to a few cells in the dorsal ectoderm (data not shown), cells in the foregut (Fig. 6n–p), surface cells in the ventral neuroectoderm (Fig. 6n, p), and a ventrolateral row of ectodermal cells (Fig. 6n, white arrowheads). Expression in the ventral neuroectoderm is a salt-and-pepper pattern, with highly expressing and weakly expressing cells interspersed with cells that do have expression (Fig. 6n). By stage 6, Ct-ngn expression in the episphere begins to decrease and is localized to a small region of anterior neuroectoderm (Fig. 6q, r, t, u). In the ventral neuroectoderm, expression decreases from anterior to posterior during stage 6, similar to the description of Ct-ash1 below. However, the decrease in Ct-ngn expression appears to precede that of Ct-ash1 (data not shown). Within the ventral neuroectoderm, in the regions where Ct-ngn expression has decreased, expression remains present in two rows of superficial cells overlying the lateral edges of the VNC (Fig. 6s, t, v, arrowheads) and in a few superficial cells near the stomodeum (Fig. 6s, t). This contrasts with expression in the more posterior ventral neuroectoderm, which has a much broader expression domain (Fig. 6s, t, u, arrow). Furthermore, expression in a small subset of cells within the VNC was detected at stage 6, but only after allowing the color reaction to proceed for a longer time. Ct-ngn also continues to be expressed in two rows of ventrolateral, ectodermal cells (Fig. 6v, white arrowhead and data not shown) and in the foregut (Fig. 6t, u). Ct-ngn expression also was faintly detected in additional cells in the ventrolateral (data not shown) and dorsolateral ectoderm (Fig. 6u; black arrowheads point to faint expression in the dorsolateral ectoderm).
From stages 7–8, Ct-ngn expression decreases considerably in most of the nervous system and is localized to superficial cells overlying the most posterior-most three to four ganglia of the VNC and to the PGZ (Additional file 7: Figure S7a–c). Within the PGZ, single Ct-ngn + cells can be detected ventrally and laterally in the ectoderm (Additional file 7: Figure S7b inset, black arrowhead). Faint, diffuse staining was also detected throughout the brain and VNC at these stages, but it was not clear whether this was actual expression because it was not ‘cellular’ in appearance. At stage 7, Ct-ngn continues to be expressed in single cells in the foregut and is expressed in a population of endodermal and possibly mesodermal cells in the trunk (Additional file 7: Figure S7a, b). By stage 9, Ct-ngn expression was only detected in the PGZ (Additional file 7: Figure S7d).
Ct-ash1 homolog expression
Ct-ash1 transcript was detected from the four-cell stage through birth of the third quartet of micromeres (data not shown). We did not examine expression in zygotes or two-cell embryos. Expression of Ct-ash1 in the brain from stages 3–6 has been previously described, so here we focus on expression in the trunk. To briefly summarize expression in the brain up to stage 6, it was found that Ct-ash1 is expressed in patches of cells in the anterior neuroectoderm starting early during gastrulation (Fig. 7a, b) . From stages 4–5, Ct-ash1 continues to be expressed in superficial cells in the anterior neuroectoderm as well as in a subset of apically localized cells in the developing brain (Fig. 7c, f) . By stage 6, Ct-ash1 is largely excluded from the brain and is expressed in individual surface cells in the anterior neuroectoderm and at the lateral edges of the brain, which may represent regions of continuing NPC ingression (Fig. 7g, j arrows).
Within the trunk at stage 4, Ct-ash1 transcripts are present in a few single cells around the stomodeum and in a circumferential ring of cells just posterior to the mouth opening (Fig. 7d, white arrowheads). By stage 5, Ct-ash1 begins to be expressed in the anterior trunk in a salt-and-pepper pattern in the ventral neuroectoderm (Fig. 7e, f) and in a subset of cells in the developing VNC . There is also expression in a cluster of cells in the pygidium (Fig. 7f, arrowhead). Expression around the stomodeum (Fig. 7e, arrows) and in the foregut also expands at this stage. By stage 6, more Ct-ash1 + cells are detected in the foregut (Fig. 7h–j), ventral neuroectoderm (Fig. 7h, j, white arrows), which continues to have a salt-and-pepper pattern (Fig. 7h), and the VNC (Fig. 7h, j, black arrowheads). At the beginning of stage 6, Ct-ash1 expression is localized to the anterior half of the neuroectoderm in the trunk (similar to Fig. 7e). As stage 6 progresses, this expression domain expands farther toward the posterior until it reaches the telotroch (data not shown). Then, toward the end of stage 6, the anterior-most limit of neuroectodermal expression in the trunk begins to regress posteriorly (Fig. 7h, i) such that expression appears to move from anterior to posterior. It was previously shown that neurons in the VNC form from anterior to posterior , and expression of Ct-ash1 in the ventral neuroectoderm may prefigure this neuronal differentiation. In the pygidium, the domain of Ct-ash1 expression expands anteriorly (Fig. 7i, j, arrowhead) , which may represent a subset of visceral mesodermal precursor cells.
At stage 7, Ct-ash1 expression in the episphere is localized to a few, single cells located in the epidermis (Fig. 7k, arrows) and at the lateral edges of the brain (data not shown). Within the trunk at stage 7, Ct-ash1 is restricted to the foregut, the presumptive posterior visceral mesoderm, and the PGZ, where neurogenesis is likely occurring (data not shown and Fig. 7k; black arrowhead points to PGZ and inset shows expression the presumptive visceral mesoderm). By stage 8, Ct-ash1 was detected faintly in the brain, while in the trunk, it was found to be localized to the pharynx and ventral portion of the esophagus, posterior visceral mesodermal cells, and the PGZ (data not shown and Additional file 7: Figure S7e; black arrowhead points to PGZ). This expression pattern is largely the same at stage 9. (Additional file 7: Figure S7f). Expression in the visceral mesoderm at stage 9 is immediately adjacent to the midgut epithelium and extends from the posterior end of the larvae anteriorly to the esophagus (Additional file 7: Figure S7f, arrowheads).
Ct-neuroD homolog expression
Ct-neuroD transcripts were first detected during early epiboly, in a few cells on the animal side of the embryo (data not shown). Later during stage 3, once a blastopore becomes apparent, Ct-neuroD is expressed in a few (approximately three to five) small, single cells in the anterior neuroectoderm (Fig. 8a–c). At this stage, Ct-neuroD is also expressed in a few cells in the lateral ectoderm (Fig. 8b, arrowheads), two cells on either side of the presumptive stomodeum (Fig. 8c, white arrowhead), and one to two cells in the presumptive pygidium (Fig. 8c, black arrowhead). At stage 4, more subsurface cells in the anterior neuroectoderm (Fig. 8d, f) as well as a few cells in the ventral neuroectoderm (Fig. 8e, arrows), and the pygidium (Fig. 8f, black arrowheads) express Ct-neuroD.
At stage 5, as the brain lobes begin to be visible, Ct-neuroD expression is limited to subsurface cells in the forming brain (Fig. 8g, i, j). In the trunk, Ct-neuroD is present in the forming ganglia of the VNC (Fig. 8h, j). During stage 6, in the head, Ct-neuroD expression is localized to basal cells within the brain lobes (Fig. 8k, m, n, o), and a few cells in the epidermis overlying the brain (Fig. 8m–o, black arrowheads). Within the trunk at stage 6, Ct-neuroD expression is maintained in all of the forming ganglia in the VNC (Fig. 8l, n, o) and is visible in a few cells in the foregut (Fig. 8n, o, arrowheads).
At stages 7 and 8, expression was no longer detected in the brain (data not shown and Additional file 7: Figure S7h, i), and only a few Ct-neuroD + cells in the more mature VNC were identified (Additional file 7: Figure S7g, i, white arrowheads). Ct-neuroD is expressed throughout the recently formed, posterior-most ganglia (Additional file 7: Figure S7g–i). Ct-neuroD was detected in a few cells in the foregut and posteriorly localized cells at the interface between the mesoderm and endoderm (Additional file 7: Figure S7h, i; arrowheads point to the pharyngeal staining). In stage 9 larvae, Ct-neuroD was only detected in the posterior-most ganglion (Additional file 7: Figure S7j).
In this article, we characterize expression of the neurogenic homologs Ct-soxB1, Ct-soxB, Ct-msi, Ct-pros, Ct-ash1, Ct-ngn, and Ct-neuroD in the annelid C. teleta. Based on their spatiotemporal patterns of expression, it is likely that many of these genes are part of the neurogenic gene regulatory network that controls development of the CNS in C. teleta.
Comparison among SoxB and bHLH group A family members in the developing CNS of Capitella teleta
Ct-soxB1, Ct-soxB, Ct-ngn, and Ct-ash1 are all expressed very early in the anterior and ventral neuroectoderm and are the earliest genes in this study to be expressed in the anterior neuroectoderm. Because expression of these genes precedes ingression of brain NPCs , this suggests a possible role in early neurogenesis such as in the maintenance of brain NPCs. Interestingly, during epiboly, the pattern of Ct-ngn differs from the other three expressed genes. Ct-ngn is expressed in two patches, each consisting of one to two large cells, at the lateral edges of the anterior ectodermal thickening (Fig. 6a, b, arrows). In contrast, Ct-soxB1, Ct-soxB, and Ct-ash1 are expressed in a medial patch of several cells (Fig. 2b, c, arrow; Fig. 3b, c, arrow; Fig. 7b). Another difference is that both Ct-soxB1 and Ct-soxB are expressed in cells outside of the anterior neuroectoderm, unlike Ct-ngn and Ct-ash1, which appear to be restricted to the neuroectoderm. Once cells start to ingress, both Ct-ngn and Ct-ash1 are expressed at varying levels in patches of surface cells, where cells divisions are occurring (see Fig. 7c, g as an example). Ct-ngn is largely restricted to surface cells, while Ct-ash1 is also expressed in a few basally localized cells. This pattern is consistent with a proneural function for both genes (see below). Ct-soxB and Ct-soxB1 continue to be expressed in neural and non-neural ectoderm in the episphere while NPCs are ingressing (i.e., at stage 4) and appear to be downregulated in the non-neural ectoderm beginning at stage 5, once fewer cells are ingressing (compare Fig. 2e with 2i). Interestingly, these genes are not restricted to the surface, where early neurogenic events occur. Instead, they are expressed throughout much of the developing brain, indicating that they may play multiple roles in neurogenesis, from NPC maintenance to neuronal commitment and specification of neuronal subtypes.
Ct-neuroD is also expressed early during gastrulation in the anterior neuroectoderm and slightly later in the ventral neuroectoderm; however, Ct-neuroD expression is entirely basal within the developing brain and VNC (e.g., Fig. 8f, j), similar to expression of the pan-neuronal genes Ct-elav and Ct-syt1 [50, 52]. The onset of Ct-neuroD expression in the anterior and ventral neuroectoderm is similar to that of Ct-ash1 and appears to precede Ct-elav and Ct-syt1. Within the developing VNC, the pattern of Ct-neuroD and Ct-elav1 is very similar during stage 6 (Fig. 9c, d). These data suggest that Ct-neuroD functions in neural cells that have exited the cell cycle, and is consistent with our previous findings that there are few dividing subsurface cells .
Comparison of Ct-soxB1 and Ct-soxB expression patterns with other metazoans
In several vertebrates, SoxB1 homologs (Sox1, 2, and 3) are expressed in overlapping patterns in the developing CNS. Expression of these homologs begins prior to formation of the neuroectoderm and then becomes restricted to proliferating NPCs. As neural daughter cells begin to differentiate, expression of SoxB1 homologs becomes downregulated, although expression is maintained in a small subset of mature neurons in the CNS [12, 14–16, 20]. SoxB1 proteins (e.g., Sox2) function primarily in dividing NPCs, while SoxB2 proteins (e.g., Sox14 and 21) function in differentiating neural cells, and antagonize SoxB1 proteins [12, 13, 17, 18, 20, 22, 93–96]. In contrast to vertebrates, genome-wide transcription factor binding studies in D. melanogaster found that the SoxB proteins SoxNeuro and Dichaete function throughout neurogenesis, from neural fate specification to neuronal maturation, and likely upregulate many of the same target genes [26, 33].
Capitella teleta Ct-soxB1 and soxB show similarities in expression with both insect and vertebrate homologs. Similar to vertebrates, Ct-soxB1 and Ct-soxB are initially expressed more broadly in the ectoderm and then become restricted to the neuroectoderm, where NPCs are actively dividing. However, both Ct-soxB1 and Ct-soxB also appear to play a role in maturing neurons, similar to insects, as expression in the brain and VNC is prominent. Furthermore, Ct-soxB expression largely overlaps that of Ct-soxB1 in the developing CNS. One interpretation is that the two homologs may be similar to insects and have redundant neurogenic functions. Alternatively, the two SoxB homologs in C. teleta could have antagonistic functions, similar to vertebrate SoxB1 and SoxB2 proteins. Furthermore, based on co-expression with Ct-ngn and Ct-ash1, the Ct-SoxB proteins may not antagonize proneural proteins as they do in vertebrates, and may instead upregulate proneural gene expression as in D. melanogaster. Interestingly, in another annelid, P. dumerilii, at least one SoxB homolog has been identified, Pdu-soxB, and it is expressed early in the neuroectoderm with subsequent downregulation upon onset of proneural gene (Pdu-neurogenin and Pdu-achaete–scute) expression [48, 49]. This pattern of expression is similar to SoxB1 expression in vertebrates and contrasts with co-expression of the soxB and proneural genes in C. teleta. Finally, to gain a better understanding of the ancestral function of SoxB proteins in spiralians, it will be important to elucidate the neurogenic gene regulatory network in C. teleta for comparison with other spiralians.
Recent work examining expression and function of a SoxB ortholog, NvSoxB(2), in the cnidarian sea anemone Nematostella vectensis indicates a role in proliferating NPCs as well as in neural cells that are not dividing [97, 98]. NvSoxB(2)+ neural cells that are dividing co-express the bHLH gene Nvath-like, while those that have exited the cell cycle co-express the proneural gene NvashA . This indicates that the interaction between SoxB homologs and bHLH type A family members in cnidarians may be more complex than a simple antagonistic relationship. Taken together, a picture is beginning to emerge in which SoxB homologs may have functioned at multiple stages of neurogenesis in the last common ancestor of cnidarians and bilaterians, a role that is supported by our data from C. teleta.
Comparison of Ct-ash1 and Ct-ngn expression patterns with other metazoans
Proneural bHLH transcription factors are key neurogenic factors in all bilaterian clades investigated so far . Proneural genes are expressed and function during neurogenesis in cnidarians [99–105], indicating an ancestral function in this process. In contrast, in ctenophores, homologs of several genes controlling neuronal fate and patterning including Neurogenin, Achaete–Scute complex, and NeuroD are absent , making it difficult to infer what molecular mechanisms governed the progression from diffuse nervous systems to more centralized systems. In vertebrates, the traditional view of proneural proteins has been that they promote cell cycle exit, neuronal differentiation, cell migration, and specification of neuronal subtypes [9, 12, 13, 18–21, 31, 107, 108], which is similar to the proposed function of NvAshA in N. vectensis. In non-insect arthropods (e.g., chelicerates and myriapods), Achaete–Scute complex homologs are thought to promote neural differentiation since they are expressed in non-dividing neural cells [39–43, 109]. This is somewhat different from expression of proneural proteins in the proliferative NPCs of vertebrates and insects [9, 44, 45].
More recently, a genome-wide transcription factor binding study in mouse found that Ascl1 (Achaete–Scute-like 1) regulates genes involved in all phases of neurogenesis [108, 110, 111]. Furthermore, there was strong evidence that Ascl1 directly promotes proliferation in brain neural progenitors. Similar genome-wide studies have not yet been conducted for vertebrate Ngn homologs, making it unclear whether they also have the ability to promote cell proliferation . Interestingly, in D. melanogaster, a genome-wide transcription factor binding study of Asense found dual functions in promoting proliferation of neuroblasts and differentiation of the ganglion mother cells .
In C. teleta, expression of Ct-ngn and Ct-ash1 in surface cells in the anterior and ventral neuroectoderm is consistent with a function in proliferating NPCs. The ‘salt-and-pepper’ expression pattern of for both genes is very similar to that seen for other proneural genes in D. melanogaster, indicating that Ct-ash1 + and Ct-ngn + cells may be undergoing lateral inhibition [9, 113]. However, whether the specification of these cells is mediated by Notch signaling still needs to be investigated. Furthermore, based on the early onset and pattern of Ct-ngn expression, we think that Ct-Ngn may play a similar role in promoting cell cycle progression as does mouse Ascl1 and insect Asense. In contrast, Ct-ash1 expression begins slightly later than Ct-ngn in the neuroectoderm and is downregulated later in the ventral neuroectoderm than Ct-ngn, indicating that Ct-Ash1 could promote cell cycle exit. This segregation of functions could be similar to that proposed for Nvath-like (dividing NPCs) and Nvash1 (differentiating neural cells) in N. vectensis [98, 104, 105]. However, preliminary experiments indicate that Ct-ash1 is expressed in both EdU+and EdU− cells (NPM unpublished data). Similarly, in P. dumerilii, Pdu-ash and Pdu-ngn are expressed apically within the ventral neuroectoderm , in the same location as proliferating cells . In addition, Pdu-ngn expression begins early in development, overlaps with Pdu-soxB, and has a widespread salt-and-pepper pattern throughout the anterior and ventral neuroectoderm, leading Simionato et al. to suggest that Pdu-Ngn could be the ‘major proneural’ factor in Platynereis . This is similar to the expression pattern we have observed for Ct-ngn; however, Ct-ngn is also expressed later in a subset of cells in the VNC. Pdu-ash expression, while apically localized, is restricted to a subset of cells in the anterior and ventral neuroectoderm, suggesting that Pdu-Ash may specify neural subtypes . In contrast, Ct-ash1 is clearly expressed in both surface and subsurface cells.
Ct-neuroD may function in neural differentiation in C. teleta
In mouse and Xenopus laevis, NeuroD homologs are expressed later than Ngn1 and 2 and primarily in post-mitotic neurons in the intermediate zone of the neural tube. Furthermore, NeuroD homologs are known to play a major role in neuronal differentiation and are direct transcriptional targets of Ngn 2 [31, 90, 107, 108, 115, 116]. NeuroD homologs are absent from D. melanogaster and Ciona intestinalis genomes, indicating lineage-specific losses . In the annelids P. dumerilii and C. teleta, Pdu-neuroD and Ct-neuroD are expressed early in the neuroectoderm, unlike in vertebrates. However, Ct-neuroD is expressed in basally localized cells within the developing brain and VNC, while Pdu-neuroD is expressed more apically and overlaps with Pdu-soxB . The expression of Ct-neuroD is consistent with a function in promoting neuronal differentiation. As NeuroD homologs have not been identified in early branching metazoans and are missing in certain taxa , the ancestral role of NeuroD is during bilaterian neurogenesis is unclear.
Ct-msi functions in post-mitotic neural cells in C. teleta
Musashi homologs are RNA-binding proteins with functions in stem-cell self-renewal, NPC maintenance, and asymmetric cell divisions in mammals and D. melanogaster [73–76, 78, 79, 81, 117]. In D. melanogaster, Musashi is necessary for asymmetric division of sensory organ precursor (SOP) cells and differentiation of their neural daughters [75, 79, 80]. In addition, msi is localized to dividing neuroblasts during larval neurogenesis in D. melanogaster , and overexpression in the larval brain induced proliferation of undifferentiated neuroblasts . The two Musashi paralogs in vertebrates, Msi-1 and Msi-2, are expressed in dividing NPCs [76, 78, 117–121], and in other chordates, Msi homologs are downregulated as the neural tube closes. Mouse Msi-2 is also widely expressed outside the CNS and in post-mitotic neurons.
Within spiralians, Msi homologs are expressed in the developing nervous system. In the planarian Dugesia japonica, expression of msi homologs is restricted to neuronal lineages, suggesting a role in neuronal differentiation . In the annelid P. dumerilii, Pdu-msi is localized to the developing brain and VNC as well as the PGZ, where cells are actively dividing . In C. teleta, Ct-msi was largely detected within the developing brain and VNC, and not in the overlying neuroectoderm, where most cell division occurs. Unlike in vertebrates, where Msi-1 homologs are downregulated in differentiating neurons, in C. teleta, Ct-msi expression is largely restricted to cells contained within the developing brain and the VNC, where NPCs are undergoing differentiation. Furthermore, at later larval stages, once neurogenesis is likely complete in the brain and anterior regions of the VNC, Ct-msi continues to be broadly expressed in the brain and VNC and is absent from the PGZ. We interpret this expression to mean that Ct-Msi is not involved in NPC maintenance or in the earlier steps of neurogenesis.
Ct-pros may be involved in asymmetric cell division of neural and non-neural ectodermal cells in C. teleta
Prospero (Pros) homologs encode homeodomain proteins that are predominantly expressed in neuronal lineages [83, 88, 123]. In both vertebrates and D. melanogaster, Prospero homologs regulate the balance between self-renewal and differentiation of neural stem cells [83, 84, 88, 112, 123–128]. In D. melanogaster, Prospero is pan-neuronal and the protein and mRNA are asymmetrically localized to the basal cortex of dividing neuroblasts and then to the basal GMC daughter cell after division. In GMC cells, Prospero enters the nucleus and induces cell cycle exit by antagonizing asense activity [83, 85, 86, 123, 127, 129, 130]. In the crustacean Daphnia magna, a prospero homolog was also found to be localized to the neuroblasts . In the spider C. salei, clusters of cells expressing an achaete–scute homolog invaginate to form the VNC. These cells also express a prospero homolog throughout the process of invagination and neuronal differentiation .
In C. teleta, Ct-pros expression suggests a role in dividing NPCs, possibly in asymmetrically dividing cells. Ct-pros is detected in single, apical cells in the anterior and ventral neuroectoderm at similar stages as Ct-ash1 and Ct-ngn. However, unlike Ct-ash1 and Ct-ngn, Ct-pros also is expressed in many ectodermal cells outside of the anterior and ventral neuroectodermal domains, suggesting an additional function in non-neural ectoderm. Ct-pros also is found in a subset of cells within the brain and VNC, and expression in the VNC persists through stage 9, long after Ct-ash1 and Ct-ngn are no longer expressed in the trunk. This suggests that Ct-pros may have additional functions in non-neural ectoderm and in neuronal subtypes. In contrast to the expression of Ct-pros in surface neuroectoderm, the P. dumerilii homolog, Pdu-prox, is expressed in groups of post-mitotic cells within the developing VNC, suggesting a function in differentiating neurons .
The expression of Ct-pros in single cells in the non-neural ectoderm of the episphere and trunk suggests that it may play a role in asymmetric division of these cells. Interestingly, one of these non-neural domains is the peristomium, where Ct-pros is detected in a circumferential band of ectodermal cells at stages 3–5 (Fig. 5d, f, k). The cells of the peristomium are morphologically distinct from other ectodermal cells in C. teleta , and to our knowledge, this is the first identified molecular marker that is predominantly expressed in the peristomial region of the trunk, if only for a short time. Ct-pros expression expands to include cells throughout the rest of the trunk ectoderm, which is most evident at stage 6 (Fig. 5o, p), indicating that the peristomial cells may initiate asymmetric cell division before the rest of the trunk. Alternatively, Ct-pros may function with other genes to specify the fate of cells in the peristomium.
We have described the spatiotemporal expression patterns of genes in C. teleta whose homologs are known to have significant functions during neurogenesis across Metazoa. Based on the expression patterns reported here, we predict that all genes examined in this study are involved in a neurogenic gene regulatory network in C. teleta. Ct-soxB1 and Ct-soxB are two of the earliest genes to be expressed, and their protein products may act to specify the neuroectoderm. Once the neuroectoderm has formed, we predict that Ct-SoxB1, Ct-SoxB, and Ct-Ngn function in dividing NPCs, possibly maintaining them in a proliferative state. Based on previous work , we think that it is likely that NPCs in C. teleta undergo asymmetric cell division in the surface neuroectoderm to generate daughters that will ingress and form the neurons of the brain and VNC. Ct-Pros may control this asymmetric cell division, while Ct-Ash1 may act in dividing NPCs to induce cell cycle exit and ingression. Ct-NeuroD likely functions in post-mitotic neural cells to direct neuronal differentiation. Finally, we think that all of the homologs examined in this study (Ct-SoxB1, Ct-SoxB, Ct-Ngn, Ct-Ash1, Ct-Pros, Ct-NeuroD, and Ct-Msi) play additional roles in post-mitotic neurons, although to varying degrees. Overall, spatiotemporal regulation of these genes in C. teleta has highlighted striking similarities as well as differences between certain aspects of neurogenesis as compared to other bilaterian clades, including variation even within annelids. Further validation of the function of these genes in C. teleta as well as continued comparative analyses is important to understand evolution of different nervous system architectures and their underlying developmental mechanisms. Our results add support to the idea of a common genetic toolkit controlling nervous system development in various cnidarian and bilaterian animals, although the molecular makeup of this toolkit has been rearranged to varying degrees during evolution, both within and across clades.
All genes were cloned by NPM except for Ct-msi, which was cloned by Danielle De Jong and Trey de Leon in the ECS Lab. WMISH for Ct-soxB1, Ct-soxB, Ct-pros, Ct-ash1, Ct-ngn, and Ct-neuroD were conducted by NPM in the in the ECS Lab. Replicate WMISH experiments for Ct-soxB, Ct-pros, and Ct-neuroD as well as multiple Ct-msi WMISH experiments were conducted by AS in the NPM Lab. Ct-soxB, Ct-pros, Ct-neuroD, and Ct-msi were imaged by AS. Ct-soxB1, Ct-ash1, Ct-ngn, and Ct-neuroD were imaged by NPM. C. teleta and Lottia gigantea Sox and Musashi gene orthologs were identified by NPM. Gene orthology analyses were conducted by CRM. Image editing and preparation of the figures were carried out by AS. AS and NPM wrote the original manuscript, and ECS and CRM gave valuable comments for improvement. All authors read and approved the final manuscript.
We thank Danielle De Jong and Trey de Leon for cloning the Ct-msi gene fragment.
The authors declare that they have no competing interests.
Availability of data and materials
Sequence data for each of the genes examined in this study have been deposited in GenBank, and accession codes will be made available upon acceptance of this manuscript. The authors declare that all other data supporting the findings of this study are available within the article and its supplementary information files.
This work was funded by the National Science Foundation (IOS09-23754 to ECS).
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- Brand AH, Livesey FJ. Neural stem cell biology in vertebrates and invertebrates: more alike than different? Neuron. 2011;70(4):719–29.PubMedView ArticleGoogle Scholar
- Hardwick LJ, et al. Cell cycle regulation of proliferation versus differentiation in the central nervous system. Cell Tissue Res. 2015;359(1):187–200.PubMedView ArticleGoogle Scholar
- Brawand D, et al. The evolution of gene expression levels in mammalian organs. Nature. 2011;478(7369):343–8.PubMedView ArticleGoogle Scholar
- Chan YF, et al. Adaptive evolution of pelvic reduction in sticklebacks by recurrent deletion of a Pitx1 enhancer. Science. 2010;327(5963):302–5.PubMedView ArticleGoogle Scholar
- Ferea TL, et al. Systematic changes in gene expression patterns following adaptive evolution in yeast. Proc Natl Acad Sci USA. 1999;96(17):9721–6.PubMedPubMed CentralView ArticleGoogle Scholar
- Harrison PW, Wright AE, Mank JE. The evolution of gene expression and the transcriptome-phenotype relationship. Semin Cell Dev Biol. 2012;23(2):222–9.PubMedView ArticleGoogle Scholar
- Alexandrova EG. Molecular mechanisms of early neurogenesis in vertebrates. Mol Biol. 2000;34:496–507.View ArticleGoogle Scholar
- Arendt D, Nubler-Jung K. Comparison of early nerve cord development in insects and vertebrates. Development. 1999;126(11):2309–25.PubMedGoogle Scholar
- Bertrand N, Castro DS, Guillemot F. Proneural genes and the specification of neural cell types. Nat Rev Neurosci. 2002;3(7):517–30.PubMedView ArticleGoogle Scholar
- Hartenstein V, Stollewerk A. The evolution of early neurogenesis. Dev Cell. 2015;32(4):390–407.PubMedView ArticleGoogle Scholar
- Wodarz A, Huttner WB. Asymmetric cell division during neurogenesis in Drosophila and vertebrates. Mech Dev. 2003;120(11):1297–309.PubMedView ArticleGoogle Scholar
- Bylund M, et al. Vertebrate neurogenesis is counteracted by Sox1-3 activity. Nat Neurosci. 2003;6(11):1162–8.PubMedView ArticleGoogle Scholar
- Graham V, et al. Sox2 functions to maintain neural progenitor identity. Neuron. 2003;39(5):749–65.PubMedView ArticleGoogle Scholar
- Ellis P, et al. Sox2, a persistent marker for multipotential neural stem cells derived from embryonic stem cells, the embryo or the adult. Dev Neurosci. 2004;26(2–4):148–65.PubMedGoogle Scholar
- Pevny L, Placzek M. Sox genes and neural progenitor identity. Curr Opin Neurobiol. 2005;15(1):7–13.PubMedView ArticleGoogle Scholar
- Wood HB, Episkopou V. Comparative expression of the mouse Sox1, Sox2 and Sox3 genes from pre-gastrulation to early somite stages. Mech Dev. 1999;86(1–2):197–201.PubMedView ArticleGoogle Scholar
- Pevny LH, Nicolis SK. Sox2 roles in neural stem cells. Int J Biochem Cell Biol. 2010;42(3):421–4.PubMedView ArticleGoogle Scholar
- Holmberg J, et al. SoxB1 transcription factors and Notch signaling use distinct mechanisms to regulate proneural gene function and neural progenitor differentiation. Development. 2008;135(10):1843–51.PubMedView ArticleGoogle Scholar
- Cau E, Blader P. Notch activity in the nervous system: to switch or not switch? Neural Dev. 2009;4:36.PubMedPubMed CentralView ArticleGoogle Scholar
- Wegner M, Stolt CC. From stem cells to neurons and glia: a Soxist’s view of neural development. Trends Neurosci. 2005;28(11):583–8.PubMedView ArticleGoogle Scholar
- Yoon K, Gaiano N. Notch signaling in the mammalian central nervous system: insights from mouse mutants. Nat Neurosci. 2005;8(6):709–15.PubMedView ArticleGoogle Scholar
- Sandberg M, Kallstrom M, Muhr J. Sox21 promotes the progression of vertebrate neurogenesis. Nat Neurosci. 2005;8(8):995–1001.PubMedView ArticleGoogle Scholar
- Phochanukul N, Russell S. No backbone but lots of Sox: invertebrate Sox genes. Int J Biochem Cell Biol. 2010;42(3):453–64.PubMedView ArticleGoogle Scholar
- Buescher M, Hing FS, Chia W. Formation of neuroblasts in the embryonic central nervous system of Drosophila melanogaster is controlled by SoxNeuro. Development. 2002;129(18):4193–203.PubMedGoogle Scholar
- Cremazy F, Berta P, Girard F. Sox neuro, a new Drosophila Sox gene expressed in the developing central nervous system. Mech Dev. 2000;93(1–2):215–9.PubMedView ArticleGoogle Scholar
- Ferrero E, Fischer B, Russell S. SoxNeuro orchestrates central nervous system specification and differentiation in Drosophila and is only partially redundant with Dichaete. Genome Biol. 2014;15(5):R74.PubMedPubMed CentralView ArticleGoogle Scholar
- Nambu PA, Nambu JR. The Drosophila fish-hook gene encodes a HMG domain protein essential for segmentation and CNS development. Development. 1996;122(11):3467–75.PubMedGoogle Scholar
- Zhao G, Skeath JB. The Sox-domain containing gene Dichaete/fish-hook acts in concert with vnd and ind to regulate cell fate in the Drosophila neuroectoderm. Development. 2002;129(5):1165–74.PubMedGoogle Scholar
- Overton PM, et al. Evidence for differential and redundant function of the Sox genes Dichaete and SoxN during CNS development in Drosophila. Development. 2002;129(18):4219–28.PubMedGoogle Scholar
- Homem CC, Knoblich JA. Drosophila neuroblasts: a model for stem cell biology. Development. 2012;139(23):4297–310.PubMedView ArticleGoogle Scholar
- Ross SE, Greenberg ME, Stiles CD. Basic helix-loop-helix factors in cortical development. Neuron. 2003;39(1):13–25.PubMedView ArticleGoogle Scholar
- Skeath JB, Thor S. Genetic control of Drosophila nerve cord development. Curr Opin Neurobiol. 2003;13(1):8–15.PubMedView ArticleGoogle Scholar
- Aleksic J, et al. The role of Dichaete in transcriptional regulation during Drosophila embryonic development. BMC Genomics. 2013;14:861.PubMedPubMed CentralView ArticleGoogle Scholar
- Mukherjee A, et al. Maternal expression and function of the Drosophila sox gene Dichaete during oogenesis. Dev Dyn. 2006;235(10):2828–35.PubMedView ArticleGoogle Scholar
- Shen SP, Aleksic J, Russell S. Identifying targets of the Sox domain protein Dichaete in the Drosophila CNS via targeted expression of dominant negative proteins. BMC Dev Biol. 2013;13:1.PubMedPubMed CentralView ArticleGoogle Scholar
- Zhao G, et al. Linking pattern formation to cell-type specification: Dichaete and Ind directly repress achaete gene expression in the Drosophila CNS. Proc Natl Acad Sci USA. 2007;104(10):3847–52.PubMedPubMed CentralView ArticleGoogle Scholar
- McKimmie C, Woerfel G, Russell S. Conserved genomic organisation of Group B Sox genes in insects. BMC Genet. 2005;6:26.PubMedPubMed CentralView ArticleGoogle Scholar
- Zhong L, et al. Parallel expansions of Sox transcription factor group B predating the diversifications of the arthropods and jawed vertebrates. PLoS ONE. 2011;6(1):e16570.PubMedPubMed CentralView ArticleGoogle Scholar
- Dove H, Stollewerk A. Comparative analysis of neurogenesis in the myriapod Glomeris marginata (Diplopoda) suggests more similarities to chelicerates than to insects. Development. 2003;130(10):2161–71.PubMedView ArticleGoogle Scholar
- Kadner D, Stollewerk A. Neurogenesis in the chilopod Lithobius forficatus suggests more similarities to chelicerates than to insects. Dev Genes Evol. 2004;214(8):367–79.PubMedView ArticleGoogle Scholar
- Stollewerk A. Recruitment of cell groups through Delta/Notch signalling during spider neurogenesis. Development. 2002;129(23):5339–48.PubMedView ArticleGoogle Scholar
- Stollewerk A, Simpson P. Evolution of early development of the nervous system: a comparison between arthropods. BioEssays. 2005;27(9):874–83.PubMedView ArticleGoogle Scholar
- Pioro HL, Stollewerk A. The expression pattern of genes involved in early neurogenesis suggests distinct and conserved functions in the diplopod Glomeris marginata. Dev Genes Evol. 2006;216(7–8):417–30.PubMedView ArticleGoogle Scholar
- Cabrera CV, Martinez-Arias A, Bate M. The expression of three members of the achaete–scute gene complex correlates with neuroblast segregation in Drosophila. Cell. 1987;50(3):425–33.PubMedView ArticleGoogle Scholar
- Skeath JB, Carroll SB. Regulation of proneural gene expression and cell fate during neuroblast segregation in the Drosophila embryo. Development. 1992;114(4):939–46.PubMedGoogle Scholar
- Monjo F, Romero R. Embryonic development of the nervous system in the planarian Schmidtea polychroa. Dev Biol. 2015;397(2):305–19.PubMedView ArticleGoogle Scholar
- Le Gouar M, Guillou A, Vervoort M. Expression of a SoxB and a Wnt2/13 gene during the development of the mollusc Patella vulgata. Dev Genes Evol. 2004;214(5):250–6.PubMedView ArticleGoogle Scholar
- Simionato E, et al. Atonal- and achaete–scute-related genes in the annelid Platynereis dumerilii: insights into the evolution of neural basic-Helix-Loop-Helix genes. BMC Evol Biol. 2008;8:170.PubMedPubMed CentralView ArticleGoogle Scholar
- Kerner P, et al. Orthologs of key vertebrate neural genes are expressed during neurogenesis in the annelid Platynereis dumerilii. Evol Dev. 2009;11(5):513–24.PubMedView ArticleGoogle Scholar
- Meyer NP, Seaver EC. Neurogenesis in an annelid: characterization of brain neural precursors in the polychaete Capitella sp. I. Dev Biol. 2009;335(1):237–52.PubMedView ArticleGoogle Scholar
- Blake A, Grassle JP, Eckelbarger KJ. Capitella teleta, a new species designation for the opportunistic and experimental Capitella sp. I, with a review of the literature for confirmed records. Zoosymposia. 2009;2:25–53.Google Scholar
- Meyer NP, et al. Nervous system development in lecithotrophic larval and juvenile stages of the annelid Capitella teleta. Front Zool. 2015;12:15.PubMedPubMed CentralView ArticleGoogle Scholar
- Seaver EC, Thamm K, Hill SD. Growth patterns during segmentation in the two polychaete annelids, Capitella sp. I and Hydroides elegans: comparisons at distinct life history stages. Evol Dev. 2005;7(4):312–26.PubMedView ArticleGoogle Scholar
- Capella-Gutierrez S, Silla-Martinez JM, Gabaldon T. trimAl: a tool for automated alignment trimming in large-scale phylogenetic analyses. Bioinformatics. 2009;25(15):1972–3.PubMedPubMed CentralView ArticleGoogle Scholar
- Darriba D, et al. ProtTest 3: fast selection of best-fit models of protein evolution. Bioinformatics. 2011;27(8):1164–5.PubMedPubMed CentralView ArticleGoogle Scholar
- Guindon S, Gascuel O. A simple, fast, and accurate algorithm to estimate large phylogenies by maximum likelihood. Syst Biol. 2003;52(5):696–704.PubMedView ArticleGoogle Scholar
- Ronquist F, et al. MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Syst Biol. 2012;61(3):539–42.PubMedPubMed CentralView ArticleGoogle Scholar
- Le SQ, Gascuel O. An improved general amino acid replacement matrix. Mol Biol Evol. 2008;25(7):1307–20.PubMedView ArticleGoogle Scholar
- Rambaut, A. FigTree v1.4.3. http://tree.bio.ed.ac.uk/software/figtree/ (2016). Accessed February 2017.
- Stamatakis A. RAxML version 8: a tool for phylogenetic analysis and post-analysis of large phylogenies. Bioinformatics. 2014;30(9):1312–3.PubMedPubMed CentralView ArticleGoogle Scholar
- Letunic I, Bork P. Interactive tree of life (iTOL) v3: an online tool for the display and annotation of phylogenetic and other trees. Nucleic Acids Res. 2016;44(W1):W242–5.PubMedPubMed CentralView ArticleGoogle Scholar
- Seaver EC, et al. The spatial and temporal expression of Ch-en, the engrailed gene in the polychaete Chaetopterus, does not support a role in body axis segmentation. Dev Biol. 2001;236(1):195–209.PubMedView ArticleGoogle Scholar
- Pevny LH, Lovell-Badge R. Sox genes find their feet. Curr Opin Genet Dev. 1997;7(3):338–44.PubMedView ArticleGoogle Scholar
- Wright EM, Snopek B, Koopman P. Seven new members of the Sox gene family expressed during mouse development. Nucleic Acids Res. 1993;21(3):744.PubMedPubMed CentralView ArticleGoogle Scholar
- Bowles J, Schepers G, Koopman P. Phylogeny of the SOX family of developmental transcription factors based on sequence and structural indicators. Dev Biol. 2000;227(2):239–55.PubMedView ArticleGoogle Scholar
- Kirby PJ, et al. Cloning and mapping of platypus SOX2 and SOX14: insights into SOX group B evolution. Cytogenet Genome Res. 2002;98(1):96–100.PubMedView ArticleGoogle Scholar
- Uchikawa M, Kamachi Y, Kondoh H. Two distinct subgroups of Group B Sox genes for transcriptional activators and repressors: their expression during embryonic organogenesis of the chicken. Mech Dev. 1999;84(1–2):103–20.PubMedView ArticleGoogle Scholar
- Wegner M. From head to toes: the multiple facets of Sox proteins. Nucleic Acids Res. 1999;27(6):1409–20.PubMedPubMed CentralView ArticleGoogle Scholar
- Schepers GE, Teasdale RD, Koopman P. Twenty pairs of sox: extent, homology, and nomenclature of the mouse and human sox transcription factor gene families. Dev Cell. 2002;3(2):167–70.PubMedView ArticleGoogle Scholar
- Kamachi Y, Uchikawa M, Kondoh H. Pairing SOX off: with partners in the regulation of embryonic development. Trends Genet. 2000;16(4):182–7.PubMedView ArticleGoogle Scholar
- Higuchi S, et al. Expression and functional analysis of musashi-like genes in planarian CNS regeneration. Mech Dev. 2008;125(7):631–45.PubMedView ArticleGoogle Scholar
- Kanemura Y, et al. Musashi1, an evolutionarily conserved neural RNA-binding protein, is a versatile marker of human glioma cells in determining their cellular origin, malignancy, and proliferative activity. Differentiation. 2001;68(2–3):141–52.PubMedView ArticleGoogle Scholar
- MacNicol AM, et al. Neural stem and progenitor cell fate transition requires regulation of Musashi1 function. BMC Dev Biol. 2015;15:15.PubMedPubMed CentralView ArticleGoogle Scholar
- Okamoto K, et al. The active stem cell specific expression of sponge Musashi homolog EflMsiA suggests its involvement in maintaining the stem cell state. Mech Dev. 2012;129(1–4):24–37.PubMedView ArticleGoogle Scholar
- Okano H, et al. Function of RNA-binding protein Musashi-1 in stem cells. Exp Cell Res. 2005;306(2):349–56.PubMedView ArticleGoogle Scholar
- Sakakibara S, et al. Mouse-Musashi-1, a neural RNA-binding protein highly enriched in the mammalian CNS stem cell. Dev Biol. 1996;176(2):230–42.PubMedView ArticleGoogle Scholar
- Yoda A, Sawa H, Okano H. MSI-1, a neural RNA-binding protein, is involved in male mating behaviour in Caenorhabditis elegans. Genes Cells. 2000;5(11):885–95.PubMedView ArticleGoogle Scholar
- Sakakibara S, et al. RNA-binding protein Musashi2: developmentally regulated expression in neural precursor cells and subpopulations of neurons in mammalian CNS. J Neurosci. 2001;21(20):8091–107.PubMedGoogle Scholar
- Nakamura M, et al. Musashi, a neural RNA-binding protein required for Drosophila adult external sensory organ development. Neuron. 1994;13(1):67–81.PubMedView ArticleGoogle Scholar
- Okabe M, et al. Translational repression determines a neuronal potential in Drosophila asymmetric cell division. Nature. 2001;411(6833):94–8.PubMedView ArticleGoogle Scholar
- Siddall NA, et al. The RNA-binding protein Musashi is required intrinsically to maintain stem cell identity. Proc Natl Acad Sci USA. 2006;103(22):8402–7.PubMedPubMed CentralView ArticleGoogle Scholar
- Yousef MS, Matthews BW. Structural basis of Prospero-DNA interaction: implications for transcription regulation in developing cells. Structure. 2005;13(4):601–7.PubMedView ArticleGoogle Scholar
- Doe CQ, et al. The prospero gene specifies cell fates in the Drosophila central nervous system. Cell. 1991;65(3):451–64.PubMedView ArticleGoogle Scholar
- Elkouris M, et al. Sox1 maintains the undifferentiated state of cortical neural progenitor cells via the suppression of Prox1-mediated cell cycle exit and neurogenesis. Stem Cells. 2011;29(1):89–98.PubMedView ArticleGoogle Scholar
- Hirata J, et al. Asymmetric segregation of the homeodomain protein Prospero during Drosophila development. Nature. 1995;377(6550):627–30.PubMedView ArticleGoogle Scholar
- Spana EP, Doe CQ. The prospero transcription factor is asymmetrically localized to the cell cortex during neuroblast mitosis in Drosophila. Development. 1995;121(10):3187–95.PubMedGoogle Scholar
- Ungerer P, Eriksson BJ, Stollewerk A. Neurogenesis in the water flea Daphnia magna (Crustacea, Branchiopoda) suggests different mechanisms of neuroblast formation in insects and crustaceans. Dev Biol. 2011;357(1):42–52.PubMedView ArticleGoogle Scholar
- Vaessin H, et al. prospero is expressed in neuronal precursors and encodes a nuclear protein that is involved in the control of axonal outgrowth in Drosophila. Cell. 1991;67(5):941–53.PubMedView ArticleGoogle Scholar
- Weller M, Tautz D. Prospero and Snail expression during spider neurogenesis. Dev Genes Evol. 2003;213(11):554–66.PubMedView ArticleGoogle Scholar
- Ledent V, Paquet O, Vervoort M. Phylogenetic analysis of the human basic helix-loop-helix proteins. Genome Biol. 2002;3(6):RESEARCH0030.PubMedPubMed CentralView ArticleGoogle Scholar
- Simionato E, et al. Origin and diversification of the basic helix-loop-helix gene family in metazoans: insights from comparative genomics. BMC Evol Biol. 2007;7:33.PubMedPubMed CentralView ArticleGoogle Scholar
- Murre C, McCaw PS, Baltimore D. A new DNA binding and dimerization motif in immunoglobulin enhancer binding, daughterless, MyoD, and myc proteins. Cell. 1989;56(5):777–83.PubMedView ArticleGoogle Scholar
- Bergsland M, et al. Sequentially acting Sox transcription factors in neural lineage development. Genes Dev. 2011;25(23):2453–64.PubMedPubMed CentralView ArticleGoogle Scholar
- Kondoh H, Kamachi Y. SOX-partner code for cell specification: regulatory target selection and underlying molecular mechanisms. Int J Biochem Cell Biol. 2010;42(3):391–9.PubMedView ArticleGoogle Scholar
- Wegner M. SOX after SOX: SOXession regulates neurogenesis. Genes Dev. 2011;25(23):2423–8.PubMedPubMed CentralView ArticleGoogle Scholar
- Sellers K, et al. Transcriptional control of GABAergic neuronal subtype identity in the thalamus. Neural Dev. 2014;9:14.PubMedPubMed CentralView ArticleGoogle Scholar
- Richards GS, Rentzsch F. Transgenic analysis of a SoxB gene reveals neural progenitor cells in the cnidarian Nematostella vectensis. Development. 2014;141(24):4681–9.PubMedView ArticleGoogle Scholar
- Richards GS, Rentzsch F. Regulation of Nematostella neural progenitors by SoxB, Notch and bHLH genes. Development. 2015;142(19):3332–42.PubMedPubMed CentralView ArticleGoogle Scholar
- Hayakawa E, Fujisawa C, Fujisawa T. Involvement of Hydra achaete–scute gene CnASH in the differentiation pathway of sensory neurons in the tentacles. Dev Genes Evol. 2004;214(10):486–92.PubMedGoogle Scholar
- Lindgens D, Holstein TW, Technau U. Hyzic, the Hydra homolog of the zic/odd-paired gene, is involved in the early specification of the sensory nematocytes. Development. 2004;131(1):191–201.PubMedView ArticleGoogle Scholar
- Grens A, et al. Evolutionary conservation of a cell fate specification gene: the Hydra achaete–scute homolog has proneural activity in Drosophila. Development. 1995;121(12):4027–35.PubMedGoogle Scholar
- Muller P, et al. Evolutionary aspects of developmentally regulated helix-loop-helix transcription factors in striated muscle of jellyfish. Dev Biol. 2003;255(2):216–29.PubMedView ArticleGoogle Scholar
- Rentzsch F, Layden M, Manuel M. The cellular and molecular basis of cnidarian neurogenesis. Wiley Interdiscip Rev Dev Biol. 2017. doi:https://doi.org/10.1002/wdev.257.PubMedGoogle Scholar
- Layden MJ, Boekhout M, Martindale MQ. Nematostella vectensis achaete–scute homolog NvashA regulates embryonic ectodermal neurogenesis and represents an ancient component of the metazoan neural specification pathway. Development. 2012;139(5):1013–22.PubMedPubMed CentralView ArticleGoogle Scholar
- Layden MJ, et al. MAPK signaling is necessary for neurogenesis in Nematostella vectensis. BMC Biol. 2016;14:61.PubMedPubMed CentralView ArticleGoogle Scholar
- Moroz LL, et al. The ctenophore genome and the evolutionary origins of neural systems. Nature. 2014;510(7503):109–14.PubMedPubMed CentralView ArticleGoogle Scholar
- Ge W, et al. Coupling of cell migration with neurogenesis by proneural bHLH factors. Proc Natl Acad Sci USA. 2006;103(5):1319–24.PubMedPubMed CentralView ArticleGoogle Scholar
- Wilkinson G, Dennis D, Schuurmans C. Proneural genes in neocortical development. Neuroscience. 2013;253:256–73.PubMedView ArticleGoogle Scholar
- Stollewerk A, Weller M, Tautz D. Neurogenesis in the spider Cupiennius salei. Development. 2001;128(14):2673–88.PubMedGoogle Scholar
- Castro DS, Guillemot F. Old and new functions of proneural factors revealed by the genome-wide characterization of their transcriptional targets. Cell Cycle. 2011;10(23):4026–31.PubMedPubMed CentralView ArticleGoogle Scholar
- Castro DS, et al. A novel function of the proneural factor Ascl1 in progenitor proliferation identified by genome-wide characterization of its targets. Genes Dev. 2011;25(9):930–45.PubMedPubMed CentralView ArticleGoogle Scholar
- Southall TD, Brand AH. Neural stem cell transcriptional networks highlight genes essential for nervous system development. EMBO J. 2009;28(24):3799–807.PubMedPubMed CentralView ArticleGoogle Scholar
- Vervoort M, Ledent V. The evolution of the neural basic helix-loop-helix proteins. Sci World J. 2001;1:396–426.View ArticleGoogle Scholar
- Denes AS, et al. Molecular architecture of annelid nerve cord supports common origin of nervous system centralization in bilateria. Cell. 2007;129(2):277–88.PubMedView ArticleGoogle Scholar
- Seo S, et al. Neurogenin and NeuroD direct transcriptional targets and their regulatory enhancers. EMBO J. 2007;26(24):5093–108.PubMedPubMed CentralView ArticleGoogle Scholar
- Ma Q, Kintner C, Anderson DJ. Identification of neurogenin, a vertebrate neuronal determination gene. Cell. 1996;87(1):43–52.PubMedView ArticleGoogle Scholar
- Kaneko Y, et al. Musashi1: an evolutionally conserved marker for CNS progenitor cells including neural stem cells. Dev Neurosci. 2000;22(1–2):139–53.PubMedView ArticleGoogle Scholar
- Sharma P, Cline HT. Visual activity regulates neural progenitor cells in developing Xenopus CNS through musashi1. Neuron. 2010;68(3):442–55.PubMedPubMed CentralView ArticleGoogle Scholar
- Shibata S, et al. Characterization of the RNA-binding protein Musashi1 in zebrafish. Brain Res. 2012;1462:162–73.PubMedView ArticleGoogle Scholar
- Imai T, et al. The neural RNA-binding protein Musashi1 translationally regulates mammalian numb gene expression by interacting with its mRNA. Mol Cell Biol. 2001;21(12):3888–900.PubMedPubMed CentralView ArticleGoogle Scholar
- Kawashima T, et al. Expression patterns of musashi homologs of the ascidians, Halocynthia roretzi and Ciona intestinalis. Dev Genes Evol. 2000;210(3):162–5.PubMedView ArticleGoogle Scholar
- Gazave E, et al. Posterior elongation in the annelid Platynereis dumerilii involves stem cells molecularly related to primordial germ cells. Dev Biol. 2013;382(1):246–67.PubMedView ArticleGoogle Scholar
- Chu-Lagraff Q, et al. The prospero gene encodes a divergent homeodomain protein that controls neuronal identity in Drosophila. Dev Suppl. 1991;2(Suppl):79–85.PubMedGoogle Scholar
- Misra K, Gui H, Matise MP. Prox1 regulates a transitory state for interneuron neurogenesis in the spinal cord. Dev Dyn. 2008;237(2):393–402.PubMedView ArticleGoogle Scholar
- Oliver G, et al. Prox 1, a prospero-related homeobox gene expressed during mouse development. Mech Dev. 1993;44(1):3–16.PubMedView ArticleGoogle Scholar
- Torii M, et al. Transcription factors Mash-1 and Prox-1 delineate early steps in differentiation of neural stem cells in the developing central nervous system. Development. 1999;126(3):443–56.PubMedGoogle Scholar
- Li L, Vaessin H. Pan-neural Prospero terminates cell proliferation during Drosophila neurogenesis. Genes Dev. 2000;14(2):147–51.PubMedPubMed CentralGoogle Scholar
- Kaltezioti V, et al. Prox1 regulates the notch1-mediated inhibition of neurogenesis. PLoS Biol. 2010;8(12):e1000565.PubMedPubMed CentralView ArticleGoogle Scholar
- Knoblich JA, Jan LY, Jan YN. Asymmetric segregation of Numb and Prospero during cell division. Nature. 1995;377(6550):624–7.PubMedView ArticleGoogle Scholar
- Choksi SP, et al. Prospero acts as a binary switch between self-renewal and differentiation in Drosophila neural stem cells. Dev Cell. 2006;11(6):775–89.PubMedView ArticleGoogle Scholar
- Meyer NP, Seaver EC. Cell lineage and fate map of the primary somatoblast of the polychaete annelid Capitella teleta. Integr Comp Biol. 2010;50(5):756–67.PubMedView ArticleGoogle Scholar